Biological Control of Bulb Mites: Comparison
Please note this is a comparison between Version 1 by Zemek Rostislav and Version 2 by Beatrix Zheng.

Mites of the genus Rhizoglyphus (Acari: Acaridae) are serious pests of plants belonging to the orders Liliales and Asparagales such as onions, garlic, lilies, and tulips. Their control by synthetic pesticides is becoming problematic as a result of resistance development in these mites and environmental and health issues. New pest control methods thus need to be developed. Efforts to develop biological control programs for bulb mites have taken place in a number of countries. Several biocontrol agents have been tested against Rhizoglyphus spp. under laboratory and some also under field conditions. The most promising results have been obtained with acaropathogenic/entomopathogenic fungi and predatory mites as described below. Other possible prospective control agents attacking mites are viruses, bacteria, and protista, but except for some bacteria their efficacy against bulb mites has not been investigated yet.

  • Rhizoglyphus
  • biological control
  • natural enemies
  • predatory mites
  • entomopathogenic fungi
  • Metarhizium
  • nematodes
  • food web
  • multitrophic interactions
  • holistic approach
  • Liliales
  • Asparagales

1. Bacteria

The soil bacterium Bacillus thuringiensis Berliner has been proven to be effective against some mite pests [1][2][53,54]. To the ouresearchers' knowledge, there are only two reports on the efficacy of this biocontrol agent or its toxins against bulb mites. The study by Carter et al. [3][29], did not find any significant effect of B. thuringiensis Cry3Aa and Cry3Bb1 coleopteran-active delta-endotoxins on Rhizoglyphus robini (Claparède). On the other hand, a recent (2020) Chinese patent [4][30] claims genetically modified B. thuringiensis to be highly efficient for Rhizoglyphus spp. control with the example of Rhizoglyphus echinopus (Fumouze and Robin) where its population was reduced by 93.2%.
Nermuť et al. [5][31] investigated whether metabolites of nematode symbiotic bacteria of the genus Xenorhabdus sp. or Photorhabdus sp. could be suitable for R. robini control. Mortality of mites treated by culture supernatants of these bacteria varied considerably among Xenorhabdus species and strains. The most effective were strains of X. doucetiae, X. bovienii, X. griffiniae and an unidentified Xenorhabdus sp., causing mortality between 10% to almost 30%. Despite the low mortality, some bacterial strains had a repellent effect on mites [5][31].

2. Acaropathogenic and Entomopathogenic Fungi

Acaropathogenic (APFs) and entomopathogenic fungi (EPFs) are common in nature, have a cosmopolitan distribution and cause natural epizootics in populations of insects, mites or other arthropods [6][7][55,56]. Fungal pathogens are a permanent component of mite natural habitats [8][57]. The advantage of EPFs, in contrast to other biocontrol agents, is that most of them are able to persist in soils for months or even years, in the absence of arthropod hosts [9][10][11][58,59,60]. A recent (2022) study by Konopická et al. [12][61] evaluated species richness and density of these fungi in soil samples collected in onion and garlic fields. EPFs Beauveria spp., Cordyceps spp., Lecanicillium spp., Metarhizium spp. and Purpureocillium spp. were isolated. The highest density was observed in the genus Metarhizium in which the average density of colony forming units (CFU) per 1 mL of soil sample reached 1.47 × 104 while the lowest density was observed in the genus Beauveria. Interestingly, soils in the Czech Republic contained about ten times higher number of EPFs compared to Israel [12][61].
Besides Acari-specific pathogens such as Hirsutella thompsonii (Fisher) and Neozygites spp. (Entomophthorales), ‘nonspecialist’ mitosporic fungi (Hyphomycetes) such as Beauveria bassiana (Bals.-Criv.) Vuill., Metarhizium anisopliae (Metsch.) Sorokin, Cordyceps fumosorosea (Wize) Kepler, B. Shrestha and Spatafora (formerly Isaria fumosorosea), Cordyceps farinosa (Holmsk.) Kepler, B. Shrestha and Spatafora (formerly Isaria farinosa), and Lecanicillium lecanii (Zimm.) Zare and W. Gams have potential to control some mite species [13][14][15][16][28,32,34,62].
Soil is considered to be a very favorable environment for EPFs application due to the high humidity conditions necessary for spore germination and host infection. On the other hand, soil-inhabiting bulb mites might have evolved at least to some extent resistance to EPFs. Indeed, a compound named hexyl rhizoglyphinate found in R. robini cuticle was shown to possess antifungal activity [17][63]. The role of other compounds, such as the monoterpenoids robinal [18][64] and isorobinal [19][65] in adaptation of bulb mites to live next to some acaro/entomopathogenic fungi in soil environments remains to be explored.
To the ouresearchers' knowledge, only four studies have tested whether APFs or EPFs are effective in controlling bulb mites under laboratory or greenhouse conditions [14][15][20][21][32,33,34,35]. Sztejnberg et al. [21][35] reported no pathogenicity of an isolate of Hirsutella kirchneri (Rostrup) Minter, Brady and Hall, obtained from the cereal rust mite, Abacarus hystrix Nalepa (original accession number CMI 257456) against R. robini, despite several procedures attempted for mite infection (spraying, dipping, tipping fungal cultures over hosts).
Konopická et al. [14][32] evaluated the effecacy of 17 isolated and 3 reference strains of EPFs against R. robini females. Results revealed high variability in R. robini mortality among EPF species and strains. The highest efficacy against R. robini mites was found in the strain of M. anisopliae isolated from soil samples collected in the Czech Republic which caused mortality up to 99.3%, and strain of Metarhizium indigoticum (Kobayasi and Shimizu) Kepler, S.A. Rehner and Humber from Israel causing 98.3% mortality, four days from spray application. The concentration-response models indicated that the latter strain was more virulent than M. anisopliae strains. The median lethal concentration (LC50) in M. indigoticum strain was estimated as 1.01 × 104. Cordyceps fumosorosea strains did not cause mortality higher than 40%. The lowest virulence was then found in Beauveria spp. strains causing mortality of mites between 5 and 25%.
Another recent (2020) study by Ment et al. [15][34] demonstrated high efficacy of Metarhizium brunneum Petch (isolate Mb7) against R. robini, which was susceptible to directly applied Mb7 conidia. Conidia of this fungus applied in vitro at concetration 1 × 107 caused 43% and 100% mortality of mites at three- and seven-days post inoculation, respectively and the estimated LT50 value was 4.3 days. Drench application in potted onion experiments also significantly reduced bulb mite populations compared to the untreated control.
The fungus Metarhizium spp. was also effective against R. robini in the study by Ko et al. [20][33]. In total, 11 isolates were selected for further study through a re-evaluation of the pathogenicity of the isolates. Conidial suspension with a concentration 1 × 107 conidia/mL of fungi Metarhizium pinghaense Q.T. Chen and H.L. Guo (isolate 3–1–2) and M. anisopliae (isolates 3–2–2 and 4–18–3) caused more than 80% mortality after 7 days. An isolate of Metarhizium pemphigi (Driver and R.J. Milner) Kepler, S.A. Rehner and Humber (isolate 1–1–1) and two isolates of M. anisopliae (isolates 4–3–2 and 4–31–2) showed mortality of 90% or more on the fifth day and 100% mortality after seven days.

3. Entomopathogenic Nematodes

Nematodes are not common parasites of mites but some mite-parasitic species are known. For example several allantonematid nematodes use mites as definitive hosts [13][28]. Usually the host is not killed but is slowly sterilised. Since these obligate parasites have not been raised on artificial media, their usefulness as biological control agents is limited. Mites can also serve as intermediate hosts of nematode parasites of vertebrates [13][28].
On the other hand, entomopathogenic nematodes (EPNs) belonging to families Steinernematidae and Heterorhabditidae can be produced in large scale with some species commercially available and successfully applied against many pests [22][23][24][25][66,67,68,69]. The only study that has explored the potential of entomopathogenic nematodes to infect bulb mites, specifically R. robini, was published in 2019 [5][31]. In this study, the bulb mites were exposed to the infective juveniles of 20 strains of Steinernema and Heterorhabditis species applied at a dose of 300 infective juveniles per mite, and the invasion rate and mite mortality were assessed. The results showed that some EPNs, especially those with small body diameter, are able to invade and kill adult females of R. robini. The most promising species were Steinernema huense Phan, Mráček, Půža, Nermuť and Jarošová, Heterorhabditis bacteriophora Poinar and Heterorhabditis amazonensis Andaló, Nguyen and Moino, which caused mortality in R. robini up to 30%. The authors concluded that although some EPN species are able to invade and kill bulb mites, their efficacy is in general quite low and they do not seem to represent a viable option for bulb mite biocontrol as a standalone approach [5][31]. EPNs have, however, other important functions in soil as they can disseminate fungal spores [26][70], serve as prey for invertebrate predators including mites and springtails or as a host for nematode-trapping fungi, such as Orbilia oligospora (Fresen.) Baral and E. Weber, Monacrosporium eudermatum (Drechsler) Subram. and Geniculifera paucispora (R.C. Cooke) Rifai [27][71].

4. Predatory Mites

Biological control programs for bulb mites have focused on using predatory mites, mainly in the family Laelapidae (Table 1), which feed on soil-dwelling pests [28][40]. Studies prior to 1990 were limited to examination of predator behavior and their ability to feed and reproduce on a diet of bulb mites. Zedan [29][42] reported that protonymphs, deutonymphs, and adults of Gaeolaelaps aculeifer (Canestrini) feed and developed on all stages of R. echinopus. Reproductive potential of the predator was highest when it fed on adult prey, but fewer prey was consumed. Ragusa and Zedan [30][43] examined interactions between these two species collected from local populations in Italy, and found that both immature and adult G. aculeifer preferred to feed on immature rather than adult R. echinopus. In contrast to Zedan [29][42], reproductive potential was highest when predators fed on a diet of eggs and immatures of R. echinopus.
Table 1. List of biocontrol agents tested for control of bulb mites.
BacteriaBacillaceaeBacillus thuringiensis Berliner[3][4]
Morganellaceae *Xenorhabdus bovienii Akhurst and Boemare[5]
Xenorhabdus budapestensis Lengyel et al.
Xenorhabdus cabanillasii Tailliez et al.
Xenorhabdus doucetiae Tailliez et al.
Xenorhabdus griffiniae Tailliez et al.
Xenorhabdus kozodoii Tailliez et al.
Xenorhabdus magdalenensis Tailliez et al.
Xenorhabdus nematophila (Poinar and Thomas) Thomas and Poinar
Xenorhabdus poinarii (Akhurst) Akhurst and Boemare
Xenorhabdus stockiae Tailliez et al.
Xenorhabdus sp.
Photorhabdus sp.
Entomo- pathogenic fungiClavicipitaceaeMetarhizium anisopliae (Metsch.) Sorokin[14][20]
Metarhizium brunneum Petch[15]
Metarhizium indigoticum (Kobayasi and Shimizu) Kepler, S.A. Rehner and Humber[14]
Metarhizium pemphigi (Driver and R.J. Milner) Kepler, S.A. Rehner and Humber[20]
Metarhizium pinghaense Q.T. Chen and H.L. Guo[20]
CordycipitaceaeBeauveria bassiana (Bals.-Criv.) Vuill.[14]
Beauveria brongniartii (Sacc.) Petch[14]
Cordyceps fumosorosea (Wize) Kepler, B. Shrestha and Spatafora[14]
OphiocordycipitaceaeHirsutella kirchneri (Rostrup) Minter, Brady and Hall[21]

pathogenic nematodes
SteinernematidaeSteinernema carpocapsae (Weiser)[5]
Steinernema huense Phan, Mráček, Půža, Nermuť and Jarošová
Steinernema surkhetense Khatri-Chhetri, Waeyenberge, Spiridonov, Manandhar and Moens
Steinernema sp.
HeterorhabditidaeHeterorhabditis amazonensis Andaló, Nguyen and Moino[5]
Heterorhabditis bacteriophora Poinar
Heterorhabditis beicherriana Li, Liu, Nermuť, Půža and Mráček
Heterorhabditis floridensis Nguyen, Gozel, Koppenhöfer and Adams
Heterorhabditis indica Poinar et al.
Heterorhabditis taysearae Shamseldean
Heterorhabditis sp.
Predatory mitesAscidaeProtogamasellus minutus Nasr[31]
BlattisociidaeLasioseius sp.[32]
Lasioseius africanus Nasr[33]
Lasioseius allii Chant (mentioned as Lasioseius bispinosus Evans)[34]
LaelapidaeCosmolaelaps barbatus Moreira, Klompen and Moraes[28]
Cosmolaelaps jaboticabalensis Moreira, Klompen and Moraes[28]
Gaeolaelaps aculeifer (Canestrini)[29][30][34][35][36][37]
Stratiolaelaps miles (Berlese)[34]
MacrochelidaeMacrocheles embersoni Azevedo, Berto and Castilho[38]
Macrocheles muscaedomesticae (Scopoli)[38]
Macrocheles robustulus (Berlese)[38]
ParasitidaeParasitus fimetorum (Berlese)[34]
RhodacaridaeProtogamasellopsis zaheri Abo-Shnaf, Castilho and Moraes (mentioned as Protogamasellopsis posnaniensis Wisniewski and Hirschmann)[39]
* Entomopathogenic nematode-associated bacteria.
In a preliminary study to find potential predators of R. robini, Lesna et al. [34][39] demonstrated that contrary to Stratiolaelaps miles (Berlese), G. aculeifer was able to feed and reproduce on a diet of R. robini. The authors also showed that local populations of G. aculeifer often differ in their feeding preference and reproductive potential and suggested that it may be advantageous to exploit these ‘local’ strains as biological control agents. In the follow-up study [35][41] the authors showed significant suppression of R. robini abundance by G. aculeifer on lilies under constant laboratory conditions and in a large-scale experiment carried out from February to June in storage rooms. Control of the pest was affected by both the spatial scale and the structural complexity of the habitat, with bulb mite populations declining faster in less complex and smaller habitats. In a greenhouse experiment using intact bulbs grown in peat soil, Lesna et al. [37][45] reported the elimination of bulb mites when the predator:prey ratio was 3:1.
Other Mesostigmata have also been reported feeding on R. echinopus [28][32][39][37,40,47] and R. robini [31][33][34][36,38,39]. Wu et al. [32][37] reported that an unidentified species of Lasioseius (Blattisociidae) developed and reproduced by feeding on R. echinopus. Castilho et al. [39][47] demonstrated that Protogamasellopsis zaheri Abo-Shnaf, Castilho and Moraes (mentioned as Protogamasellopsis posnaniensis Wisniewski and Hirschmann) (Rhodacaridae) fed on R. echinopus and reproduced. Moreira and Moraes [28][40] observed that Cosmolaelaps barbatus Moreira, Klompen and Moraes and Cosmolaelaps jaboticabalensis Moreira, Klompen and Moraes (Laelapidae) oviposited when fed R. echinopus. Azevedo et al. [38][46] observed that Macrocheles embersoni Azevedo, Berto and Castilho, Macrocheles muscaedomesticae (Scopoli) and Macrocheles robustulus (Berlese) (Macrochelidae) were able to consume R. echinopus, but with a very low consumption compared to other prey. In the case of R. echinopus, Afifi et al. [31][36] observed the development and reproduction of Protogamasellus minutus Nasr (Ascidae) when fed with this prey. Also, Lesna et al. [34][39] reported that Lasioseius allii Chant (mentioned as Lasioseius bispinosus Evans) (Blattisociidae) and Parasitus fimetorum (Berlese) (Parasitidae) were successfully reared on R. robini, and the latter was able to suppress the prey when peat was used as substrate. Mowafi [33][38] observed the development and reproduction of Lasioseius africanus Nasr (Blattisociidae) feeding on R. robini. However, all of these studies were conducted in the laboratory and only in the case of P. fimetorum were small-scale population experiments conducted in closed flasks. Further studies are needed to explore the potential of these species as biocontrol agents in potted plant and field trials.
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